We acknowledge the Human Immunology Core for providing normal donor T cells and the Flow Cytometry Core at University of Pennsylvania.

Human T cells were procured through the University of Pennsylvania Human Immunology Core, which operates under principles of Good Laboratory Practice with established standard operating procedures and/or protocols for sample receipt, processing, freezing, and analysis conform to MIATA and University of Pennsylvania ethics guidelines.
1. Lentiviral vector production
NOTE: The viral products have been made replication-defective by separation of packaging constructs (Rev, gag/pol/RRE, VSVg and transfer plasmid) into four separate plasmids, greatly reducing the likelihood of recombination events that may result in replication-competent virus.
<ol>
	<li>Prepare HEK293T cells one day prior to the transfection. Plate approximately 6 x 106 cells in T150 culture vessels in 30 mL of standard culture media (referred to as R10:RPMI 1640 supplemented with 10% fetal calf serum, 10 mM HEPES, 1% Pen/Strep, 1% L-glutamine) and incubate overnight at 37 &#176;C. 18-24 h later, cells should be 60-70% confluent and look healthy (dendritic projections and uniform distribution across the flask). If cells look healthy, proceed to step 1.2.</li>
	<li>Prepare the transfection mix containing lipofection reagent (96 &#181;L), pTRP gag/pol (Lot# RR13SEP19A) (18 &#181;g), pTRP RSV-Rev (Lot# RR13SEP19B-3) (18 &#181;g), pTRP VSVG (Lot# RR13SEP19C) (7 &#181;g) packaging plasmids and 15 &#181;g of expression plasmid (CD19bbz scFv cloned in pTRPE).
	<ol>
		<li>For each transfection reaction, prepare one tube containing 1 mL of reduced-serum minimal essential media plus the four plasmids described in step 1.2 as well as one tube containing 2 mL of reduced-serum minimal essential media plus 90 &#181;L of lipofection reagent. Combine these two solutions by dropwise addition of the 1 mL plasmid mix to the 2 mL of lipofection reagent mix. Be sure to vigorously mix the solution by pipetting up and down several times. Incubate the solution for 15 min at room temperature.</li>
		<li>In the meantime, aspirate the media from the HEK293T cell flask and gently wash cells with 10 mL of reduced-serum minimal essential media and aspirate again.</li>
		<li>Gently add the transfection mix (3 mL) from step 1.2.1 to the bottom corner of cell culture flask using a 5 mL serological pipette. Gently tilt the flask to evenly distribute the transfection mix across the cell culture flask and incubate for 10 min at room temperature. Make sure to not disturb the cell monolayer. After a 10 min incubation, add 35 mL of R10 media and return flask to the incubator for 24 h.</li>
	</ol>
	</li>
	<li>After 24 h, collect the supernatant from the HEK293T cell culture flasks and transfer into 50 mL conical tubes. Add fresh R10 to the HEK29T cells and put the cells back into the incubator for another 24 h. Spin down the supernatant (300 x g) to remove cell debris and filter the supernatant through a 0.45 &#181;m filter.</li>
	<li>Concentrate the filtrate from step 1.3 by high-speed ultracentrifugation (25,000 x g for 2.5 h or 8000 x g overnight (O/N)). Store the 24 h virus pellet at 4 &#176;C while pooling the 48 h virus.</li>
	<li>Repeat steps 1.3-1.4 for the 48 h collection and combine 24 h and 48 h virus collections. Pellets from the 48 h collection will contain a combined harvest of the lentivirus.</li>
	<li>Resuspend viral pellet in approximately 1 mL of cold R10 and aliquot into 100 &#181;L vials. Immediately snap freeze the aliquots using dry ice and store at -80 &#176;C.</li>
	<li>Calculate functional viral titer in transducing units/mL by transducing a fixed amount of CD3/CD28 bead-activated primary human T-cells with serial dilutions of the lentiviral supernatant. Measure CAR-Transduction after 72 h by flow-cytometry.
	<ol>
		<li>Stain for the CD19-directed CAR with an anti-FCM63 scFv antibody. For other chimeric antigen receptors, stain using fluorophore-labeled recombinant protein that is specific to the CAR scFv. Virus dilution that generates under 20% CAR positive cells by flow-cytometry is the most accurate dilution to calculate viral titer from. Above 20% CAR-positive cells, the chance for each positive cell to be transduced twice increases, resulting in an underestimation of the number of transducing particles. Calculate the transducing units per mL(TU/mL ) = (number of cells transduced x percent CAR positive cells x dilution factor)/(transduction volume in mL).</li>
	</ol>
	</li>
</ol>
2. Designing of sgRNAs and gene disruption in primary human T cells
<ol>
	<li>Designing CRISPR sgRNA&#39;s 
	NOTE: Several servers and programs facilitate the design of target-specific sgRNA&#39;s. In this protocol, CRISPR sgRNAs were designed using CHOPCHOP (https://chopchop.cbu.uib.no), and the gRNA design portal from the Broad institute (https://portals.broadinstitute.org/gpp/public/analysis-tools/sgrna-design).
	<ol>
		<li>For each target gene, design six to ten sgRNA sequences to target early coding exon sequences. 
		NOTE: sgRNA should have high on-target efficacy and low off target efficacy. Each machine learning algorithm for determining sgRNA efficacy works slightly differently. Therefore, comparing multiple sgRNA design tools and curating a list of six to ten sgRNA for screening is recommended.</li>
	</ol>
	</li>
	<li>Gene disruption in primary human T-cells
	<ol>
		<li>Obtain autologous peripheral blood mononuclear cells (PBMC&#39;s) from healthy volunteer donors. A schematic timeline of the protocol is described in Figure 2A and described below.</li>
		<li>Isolate CD4+ and CD8+ T-cells using commercially available CD4 and CD8 selection kits.</li>
		<li>Combine CD4+ and CD8+ T-cells at a 1:1 ratio and incubate overnight at 3x106 cells/mL in R10 supplemented with 5 ng/mL huIL-7 and huIL-15 each. Addition of IL-7 and IL-15 is recommended as they are known to promote a central memory phenotype and CAR-T cells expanded with IL-7 and IL-15 have superior anti-tumor efficacy compared to IL-2 expanded CAR-T cells22,23,24.</li>
		<li>The next day, count T-cells and centrifuge 5-10 x 106 cells at 300 x g for 5 min. Discard all the supernatant and wash the cell pellet in reduced-serum minimal essential media. Resuspend the pellet in 100 &#181;L of nucleofection solution according to the manufacturer&#39;s instructions (Table of Materials).</li>
		<li>While washing the cells, prepare ribonucleoprotein (RNP) complex with Cas9 and gRNA by incubating 10 &#181;g of Cas9 nuclease with 5 &#181;g of sgRNA for 10 min at room temperature (RT). The molar ratio of Cas9 to sgRNA is 1:2.4. A mock control that contains Cas9 and electroporation enhancer, but no sgRNA, is recommended. 
		NOTE: Multiplex gene editing can be performed at this step by making RNP complexes for each target separately and combining them with cells at the time nucleofection as described in the next step. Choosing the reagents to assemble the RNP complex are key to getting high KO efficiency. For example, Cas9 nucleases from different vendors have varying off-target effects and chemically modified gRNA decrease toxicity to T cells, thereby increasing KO efficiency.</li>
		<li>Combine the resuspended cells from step 2.2.4 with the RNP complex from step 2.2.5 and add 4.2 &#181;L of 4 &#181;M electroporation enhancer (ssDNA oligonucleotide that is non-homologous to human genome). Mix well and transfer into electroporation cuvettes. Avoid bubbles as they impair electroporation efficiency.</li>
		<li>Electroporate cells using pulse code EH111. After electroporation, incubate cells in R10 supplemented with 5 ng/mL huIL-7 and huIL-15 at 5x106 cells/mL at 30 &#176;C for 48 h in 12-well plates. After 48 h, proceed with T-cell activation and expansion.</li>
	</ol>
	</li>
</ol>
3. T cell activation, lentiviral transduction and expansion
NOTE: For screening sgRNA&#39;s, lentiviral transduction (Step 3.2) of the CAR construct is not necessary.
<ol>
	<li>Count electroporated T-cells and dilute to a concentration of 1 x 106 cells/mL using warm R10 supplemented with 5 ng/mL huIL-7 and huIL-15. Activate cells by adding anti-CD3/anti-CD28 monoclonal antibody coated magnetic beads at a ratio of 3 beads per live T-cell. 
	NOTE: Electroporation causes significant cell death resulting in roughly 60% &#177; 15% viable cells compared to non-electroporated cells. Use a live/dead cell count to determine the appropriate amount of beads. Transfer the cells to 37 &#176;C and incubate overnight.</li>
	<li>After overnight stimulation, transduce CRISPR-engineered T-cells with lentiviral supernatant from step 1. Add the appropriate volume based on the viral titer calculated in step 1.7 to achieve a multiplicity of infection (MOI) of 3 and incubate cells for 72 h at 37 &#176;C. 
	NOTE: The virus pellet resuspended in R10 is simply added on top of the cells according to the cell concentration, viral titer, and desired MOI.</li>
	<li>After 72 h, feed cells with 50% of the current culture volume R10 containing 10 ng/mL huIL-7 and huIL-15 and return them to the incubator for another 48 h. Do not disturb the clusters that have formed between the T-cells and the beads.</li>
	<li>After 48 h, remove the beads from the cells by gently resuspending the cell-bead mixture followed by magnetic separation. Count cells after bead removal and bring to a concentration of 0.8 x 106 cells/mL using R10 supplemented with 5 ng/mL huIL-7 and huIL-15. After de-beading, cell numbers should be similar to the cell count on Day 1 prior to electroporation (Figure 2B).</li>
	<li>After 24 h, count cells and feed to a concentration of 1 x 106 cells/mL with R10 + 5 ng/mL huIL-7 and huIL-15. Repeat this every 24 h until growth kinetics and cell size demonstrate cells have rested from stimulation. 
	NOTE: From this point, T-cells double approximately every 24h. T-cell usually undergo 5-7 population doublings and have a cell volume of approximately 300 &#177; 50 fL when rested.</li>
	<li>Once rested, spin down CRISPR-engineered CAR-T cells and resuspend in freeze media (1:1 X-Vivo and FBS plus 10% DMSO) for cryopreservation. CAR-T cells used should be thawed and rested at 37 &#176;C for 16 hours before an experiment. 
	NOTE: Keep approximately 10x106 cells reserved for measuring knockout efficiency and CAR integration. Expected population doublings and volume changes during the expansion have been shown in Figure 2B,2C. Additionally, Figure 2D shows levels of CAR expression on both Mock and gene edited CAR-T cells.</li>
</ol>
4. CRISPR Efficiency
<ol>
	<li>Genomic DNA extraction and target gene amplification
	<ol>
		<li>From each screening culture, spin down 3-5 x 106 T-cells. Cells can be either frozen down as a dry pellet or one can proceed with genomic DNA extraction. At time of DNA extraction, resuspend pellets in 200 &#181;L of Phosphate Buffer Saline (PBS) and extract genomic DNA from electroporated cells using a standard DNA extraction kit according to the manufacturer instruction.
		<ol>
			<li>Briefly, lyse cells with proteinase K and load lysate onto a DNA binding column. During centrifugation, DNA is binding to the membrane while contaminants will pass through. After two washing steps to remove remaining contaminants and enzyme inhibitors, elute DNA in water.</li>
		</ol>
		</li>
		<li>Amplify 200-300 ng of genomic DNA using standard PCR mix containing DNA polymerase, accessory proteins, salts, and dNTPs and 10 &#181;M forward and reverse primers flanking the region of the intended double-strand break. 
		NOTE: For PCR primer design, the reference genomic sequence (can be found using http://www.ensemble.org) flanking the gRNA cut site are entered into the NCBI primer blast tool (https://www.ncbi.nlm.nih.gov/tools/primer-blast). PCR primers should be designed such that the amplicon has a target size of 600-700bp. This length allows to design sequencing primers that bind within the amplicon with sufficient distance from the gRNA cut site (at least 150 bp) to ensure good sequencing quality around the Cas9 induced indels (see 4.2.1). Each gRNA requires a unique primer pair, unless the gRNA cut sites are in close proximity.&#160;</li>
		<li>Run the entire PCR product on a 1% agarose gel and purify the amplicon using a standard agarose gel purification kit. 
		NOTE: The process of determining the optimal Tm for PCR can take multiple rounds of troubleshooting. This is because the KO sequence has indels and the target gene should be primed for sequencing where there would be no indels usually 100 bp upstream or downstream of the cut site. This can vary widely due to indels resulting from non-homologous end joining. Designing nested sequencing primers to the PCR primers and increasing the concentration of product sequenced can eliminate problems with sequencing.</li>
	</ol>
	</li>
	<li>Sequencing and indel detection
	<ol>
		<li>Design sequencing primers that bind to the gel purified amplicon and send mock and knockout amplicons for Sanger sequencing. Once sequencing is complete, upload the trace files to the Desktop Genetics software (tide.deskgen.com, Desktop Genetics) for TIDE analysis.
		<ol>
			<li>For sequencing primer design, enter the sequence of the PCR amplicon into a standard primer design software (NCBI, Eurofins or other publicly available tools). The design software will suggest multiple primer sequences that are suitable to sanger sequencing.</li>
			<li>Choose forward and reverse primers that bind within the amplicon at least 150 bp upstream or downstream of the gRNA cut site to ensure good sequencing quality around the indels. The sequencing chromatogram will be used in 4.2 for TIDE analysis, therefore it is important that sequencing quality is good for accurate indel decomposition (see Hultquist et al.)25.</li>
		</ol>
		</li>
		<li>Use TIDE (Tracking of Indels by DEcomposition) analysis to detect knock out (KO) efficiency at the genomic level26. The algorithm accurately reconstructs the spectrum of indels from the sequence traces and calculates R2 values, reflecting goodness of fit after non-negative linear modeling by the TIDE software.</li>
	</ol>
	</li>
	<li>Target protein detection
	<ol>
		<li>For target genes whose gene product is expressed on the surface of T-cells, stain approximately 1 x 105 cells from each screening culture with a fluorochrome-tagged antibody specific for the target protein. Compare expression levels of the target protein between the mock control and knockout groups by flow cytometry as shown in Figure 3A,3B.</li>
		<li>For target genes whose gene product is expressed intracellularly, lyse approximately 3 x 106&#160;million cells per screening group in lysis buffer and follow standard protocols for SDS-PAGE and western blotting. Alternatively, surface or intracellular flow-cytometry can be used to probe for the target protein.</li>
	</ol>
	</li>
</ol>
5. Monitoring Off-target effects using iGUIDE - Library preparation, DNA sequencing, and analysis
NOTE: iGUIDE technique allows for detection of locations of Cas9 guided cleavage and quantify the distributions of those DNA double-stranded breaks.
<ol>
	<li>Perform iGUIDE as described by Nobles et al.21. The off-target effects of sgRNA targeting TRAC locus using the iGUIDE technique have been shown in Stadtmauer et al, 202013.</li>
</ol>

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The authors have no disclosures.

Here we describe approaches to gene edit CAR-T cells using CRISPR Cas9 technology and manufacture products to further test for function and efficacy. The above protocol has been optimized for performing CRIPSR gene editing in primary human T cells combined with engineering T cells with chimeric antigen receptors. This protocol allows high knockout efficiency with minimal donor-to-donor variability. Modification using CRISPR can improve both the efficacy and safety of CAR-T cells by eliminating receptors that inhibit T ce...

Chimeric antigen receptor (CAR)-T cell therapy has revolutionized the field of adoptive cell therapies and cancer immunotherapy. CAR-T-cells are engineered T-cells expressing a synthetic immune receptor that combines an antigen-specific single chain antibody fragment with signaling domains derived from the TCRzeta chain and costimulatory domains necessary and sufficient for T-cell activation and co-stimulation1,2,3,4. The manufacturing of CAR-T cells starts by extracting the patient&#39;s own T-cells, followed by ex vivo viral transduction of the CAR module and expansion of the CAR-T cell product with magnetic beads that function as artificial antigen presenting cells5. Expanded CAR-T cells are re-infused into the patient where they can engraft, eliminate target tumor cells and even persist for multiple years post infusion6,7,8. Although CAR-T cell therapy has resulted in remarkable response rates in B-cell malignancies, clinical success for solid tumors has been challenged by multiple factors including poor T-cell infiltration9, an immunosuppressive tumor microenvironment10, antigen coverage and specificity, and CAR-T cell dysfunction11,12. Another limitation of current CAR-T cell therapy includes the use of autologous T-cells. After multiple rounds of chemotherapy and high tumor burden, CAR-T cells can be of poor quality as compared to allogeneic CAR-T products from healthy donors in addition to the time and expense associated with manufacturing of autologous CAR-T cells. Gene-editing of the CAR-T cell product by CRISPR/Cas9 represents a new strategy to overcome current limitations of CAR-T cells13,14,15,16,17.
CRISPR/Cas9 is a two component system that can be used for targeted genome editing in mammalian cells18,19. The CRISPR-associated endonuclease Cas9 can induce site-specific double-strand breaks guided by small RNAs through base-pairing with the target DNA sequence20. In the absence of a repair template, double-strand breaks are repaired by the error prone nonhomologous end joining (NHEJ) pathway, resulting in frameshift mutations or premature stop codons through insertion and deletion mutations (INDELs)19,20,21. Efficiency, ease of use, cost-effectiveness and the ability for multiplex genome editing make CRISPR/Cas9 a powerful tool to enhance the efficacy and safety of autologous and allogeneic CAR-T cells. This approach can also be used to edit TCR directed T cells by replacing the CAR construct with a TCR. Additionally, allogeneic CAR-T cells that have limited potential to cause graft versus host disease can also be generated by gene editing the TCR, b2m, and HLA locus.
In this protocol, we show how CRISPR-engineering of T cells can be combined with viral vector mediated delivery of the CAR-Transgene to generate genome-edited CAR-T cell products with enhanced efficacy and safety. A schematic diagram of the entire process is shown in Figure 1. Using this approach, we have demonstrated high-efficiency gene knockout in primary human CAR-T cells. Figure 2A describes in detail the timeline of each step for editing and manufacturing T cells. Strategies for guide RNA design and knockout validation are also discussed to apply this approach to various target genes.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>4D-Nucleofactor Core Unit</td>
			<td>Lonza</td>
			<td>AAF-1002B</td>
			<td></td>
		</tr>
		<tr>
			<td>4D-Nucleofactor X-Unit</td>
			<td>Lonza</td>
			<td>AAF-1002X</td>
			<td></td>
		</tr>
		<tr>
			<td>Accuprime Pfx Supermix</td>
			<td>ThermoFisher</td>
			<td>12344040</td>
			<td></td>
		</tr>
		<tr>
			<td>Beckman Optima XPN ultracentrifuge</td>
			<td>Beckman Coulter</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Brilliant Violet 605 anti-human CD3 Antibody</td>
			<td>Biolegend</td>
			<td>317322</td>
			<td>Clone OKT3</td>
		</tr>
		<tr>
			<td>BV711 Anti-human PD1</td>
			<td>Biolegend</td>
			<td></td>
			<td>Clone EH12.2H7</td>
		</tr>
		<tr>
			<td>Cas9-Electroporation enhancers</td>
			<td>IDT</td>
			<td>1075915</td>
			<td></td>
		</tr>
		<tr>
			<td>CD3/CD28 Dynabeads</td>
			<td>ThermoFisher</td>
			<td>40203D</td>
			<td></td>
		</tr>
		<tr>
			<td>CD4+ T cell isolation Kit</td>
			<td>StemCell technologies</td>
			<td>15062</td>
			<td></td>
		</tr>
		<tr>
			<td>CD8+ T cell isolation Kit</td>
			<td>StemCell technologies</td>
			<td>15063</td>
			<td></td>
		</tr>
		<tr>
			<td>Corning 0.45 micron vacuum filter/bottle</td>
			<td>Corning</td>
			<td>430768</td>
			<td></td>
		</tr>
		<tr>
			<td>Corning T150 cell culture flask</td>
			<td>Millipore Sigma</td>
			<td>CLS430825</td>
			<td></td>
		</tr>
		<tr>
			<td>DMSO</td>
			<td>Millipore Sigma</td>
			<td>D2650</td>
			<td></td>
		</tr>
		<tr>
			<td>DNAeasy Blood and Tissue Kit</td>
			<td>Qiagen</td>
			<td>69504</td>
			<td></td>
		</tr>
		<tr>
			<td>DynaMag Magnet</td>
			<td>ThermoFisher</td>
			<td>12321D</td>
			<td></td>
		</tr>
		<tr>
			<td>Glutamax supplement</td>
			<td>ThermoFisher</td>
			<td>35050061</td>
			<td></td>
		</tr>
		<tr>
			<td>HEK293T cells</td>
			<td>ATCC</td>
			<td>CRL-3216</td>
			<td></td>
		</tr>
		<tr>
			<td>HEPES (1 M)</td>
			<td>ThermoFisher</td>
			<td>15630080</td>
			<td></td>
		</tr>
		<tr>
			<td>huIL-15</td>
			<td>PeproTech</td>
			<td>200-15</td>
			<td></td>
		</tr>
		<tr>
			<td>huIL-7</td>
			<td>PeproTech</td>
			<td>200-07</td>
			<td></td>
		</tr>
		<tr>
			<td>Lipofectamine 2000</td>
			<td>ThermoFisher</td>
			<td>11668019</td>
			<td></td>
		</tr>
		<tr>
			<td>Nucleospin Gel and PCR cleanup</td>
			<td>Takara</td>
			<td>740609.25</td>
			<td></td>
		</tr>
		<tr>
			<td>Opti-MEM</td>
			<td>ThermoFisher</td>
			<td>31985062</td>
			<td></td>
		</tr>
		<tr>
			<td>P3 Primary cell 4D-nucleofactor X Kit L</td>
			<td>Lonza</td>
			<td>V4XP-3024</td>
			<td></td>
		</tr>
		<tr>
			<td>Penicilin-Streptomycin-Glutamine</td>
			<td>ThermoFisher</td>
			<td>10378016</td>
			<td></td>
		</tr>
		<tr>
			<td>pTRPE expression Plasmid</td>
			<td>in house</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Rabbit Anti-Mouse FMC63 scFv Monoclonal Antibody, (R19M), PE</td>
			<td>CytoArt</td>
			<td>200105</td>
			<td></td>
		</tr>
		<tr>
			<td>RPMI1640</td>
			<td>ThermoFisher</td>
			<td>12633012</td>
			<td></td>
		</tr>
		<tr>
			<td>sgRNA</td>
			<td>IDT</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Spy Fi Cas9</td>
			<td>Aldevron</td>
			<td>9214</td>
			<td></td>
		</tr>
		<tr>
			<td>Ultracentrifuge tubes</td>
			<td>Beckman Coulter</td>
			<td>326823</td>
			<td></td>
		</tr>
		<tr>
			<td>Viral packaging mix</td>
			<td>in house</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>X-Vivo-15 Media</td>
			<td>Lonza</td>
			<td>BE02-060F</td>
			<td></td>
		</tr>
	</tbody>
</table>

production of human crispr-engineered car-t cells

Adoptive cell therapies using chimeric antigen receptor T cells (CAR-T cells) have demonstrated remarkable clinical efficacy in patients with hematological malignancies and are currently being investigated for various solid tumors. CAR-T cells are generated by removing T cells from a patient's blood and engineering them to express a synthetic immune receptor that redirects the T-cells to recognize and eliminate target tumor cells. Gene editing of CAR-T cells has the potential to improve safety of current CAR-T cell therapies and further increase the efficacy of CAR-T cells. Here, we describe methods for the activation, expansion, and characterization of human CRISPR-engineered CD19 directed CAR-T cells. This comprises transduction of the CAR lentiviral vector and use of single guide RNA (sgRNA) and Cas9 endonuclease to target genes of interest in T cells. The methods described in this protocol can be universally applied to other CAR constructs and target genes beyond the ones used for this study. Furthermore, this protocol discusses strategies for gRNA design, lead gRNA selection and target gene knockout validation to reproducibly achieve high-efficiency, multiplex CRISPR-Cas9 engineering of clinical grade human T cells.

We describe here a protocol to genetically engineer T cells, that can be used to generate both autologous and allogeneic CAR-T cells, as well as TCR redirected T cells.
Figure 1 provides a detailed description of the stages involved in the process of manufacturing CRISPR edited T cells. The process begins by designing sgRNA to the gene of interest. Once the sgRNA are designed and synthesized they are then used to make RNP complexes with the appropriate Cas protein...

Watch this Scientific Journal Video about Production of Human CRISPR-Engineered CAR-T Cells at JoVE.com

Production of Human CRISPR-Engineered CAR-T Cells

Here, we present a protocol for gene editing in primary human T cells using CRISPR Cas Technology to modify CAR-T cells.

Adoptive cell therapies using chimeric antigen receptor T cells (CAR-T cells) have demonstrated remarkable clinical efficacy in patients with ...

production-of-human-crispr-engineered-car-t-cells

Research

JoVE Journal

Immunology and Infection

12.4K Views.  University of Pennsylvania. Here, we present a protocol for gene editing in primary human T cells using CRISPR Cas Technology to modify CAR-T cells.

Video: Production of Human CRISPR-Engineered CAR-T Cells

Recombinant Retroviral Production and Infection of B Cells

The transgenic expression of genes in eukaryotic cells is a powerful reverse genetic approach in which a gene of interest is expressed under the control of a heterologous expression system to facilitate the analysis of the resulting phenotype. This approach can be used to express a gene that is not normally found in the organism, to express a mutant form of a gene product, or to over-express a dominant-negative form of the gene product. It is particularly useful in the study of the hematopoetic system, where transcriptional regulation is a major control mechanism in the development and differentiation of B cells 1, reviewed in 2-4.
Mouse genetics is a powerful tool for the study of human genes and diseases. A comparative analysis of the mouse and human genome reveals conservation of synteny in over 90% of the genome 5. Also, much of the technology used in mouse models is applicable to the study of human genes, for example, gene disruptions and allelic replacement 6. However, the creation of a transgenic mouse requires a great deal of resources of both a financial and technical nature. Several projects have begun to compile libraries of knock out mouse strains (KOMP, EUCOMM, NorCOMM) or mutagenesis induced strains (RIKEN), which require large-scale efforts and collaboration 7. Therefore, it is desirable to first study the phenotype of a desired gene in a cell culture model of primary cells before progressing to a mouse model. 
Retroviral DNA integrates into the host DNA, preferably within or near transcription units or CpG islands, resulting in stable and heritable expression of the packaged gene of interest while avoiding transcriptional silencing 8 9. The genes are then transcribed under the control of a high efficiency retroviral promoter, resulting in a high efficiency of transcription and protein production. Therefore, retroviral expression can be used with cells that are difficult to transfect, provided the cells are in an active state during mitosis. Because the structural genes of the virus are contained within the packaging cell line, the expression vectors used to clone the gene of interest contain no structural genes of the virus, which both eliminates the possibility of viral revertants and increases the safety of working with viral supernatants as no infectious virions are produced 10. 
Here we present a protocol for recombinant retroviral production and subsequent infection of splenic B cells. After isolation, the cultured splenic cells are stimulated with Th derived lymphokines and anti-CD40, which induces a burst of B cell proliferation and differentiation 11. This protocol is ideal for the study of events occurring late in B cell development and differentiation, as B cells are isolated from the spleen following initial hematopoetic events but prior to antigenic stimulation to induce plasmacytic differentiation.

An efficient system of structure and function analysis of a gene in an ex vivo culture of splenic B-lymphocytes is described. This method takes advantage of recombinant retroviral production in a helper free, ecotrophic packaging cell line. Stable, heritable expression of a gene of interest within primary lymphocytes is achieved leading to generation of surface antibodies on B cells undergoing class switch recombination.

The transgenic expression of genes in eukaryotic cells is a powerful reverse genetic approach in which a gene of interest is expressed under the control ...

Invasion of Human Cells by a Bacterial Pathogen

Here we will describe how we study the invasion of human endothelial cells by bacterial pathogen Staphylococcus aureus . The general protocol can be applied to the study of cell invasion by virtually any culturable bacterium. The stages at which specific aspects of invasion can be studied, such as the role of actin rearrangement or caveolae, will be highlighted. Host cells are grown in flasks and when ready for use are seeded into 24-well plates containing Thermanox coverslips. Using coverslips allows subsequent removal of the cells from the wells to reduce interference from serum proteins deposited onto the sides of the wells (to which S. aureus would attach). Bacteria are grown to the required density and washed to remove any secreted proteins (e.g. toxins). Coverslips with confluent layers of endothelial cells are transferred to new 24-well plates containing fresh culture medium before the addition of bacteria. Bacteria and cells are then incubated together for the required amount of time in 5% CO2 at 37&deg;C. For S. aureus this is typically between 15-90 minutes. Thermanox coverslips are removed from each well and dip-washed in PBS to remove unattached bacteria. If total associated bacteria (adherent and internalised) are to be quantified, coverslips are then placed in a fresh well containing 0.5% Triton X-100 in PBS. Gentle pipetting leads to complete cell lysis and bacteria are enumerated by serial dilution and plating onto agar. If the number of bacteria that have invaded the cells is needed, coverslips are added to wells containing 500 &mu;l tissue culture medium supplemented with gentamicin and incubation continued for 1 h, which will kill all external bacteria. Coverslips can then be washed, cells lysed and bacteria enumerated by plating onto agar as described above. If the experiment requires direct visualisation, coverslips can be fixed and stained for light, fluorescence or confocal microscopy or prepared for electron microscopy.

A general protocol for the study of invasion of host cells by a bacterial pathogen, focusing on Staphylococcus aureus and human endothelial cells.

Here we will describe how we study the invasion of human endothelial cells by bacterial pathogen Staphylococcus aureus . The general protocol can be ...

piggyBac Transposon System Modification of Primary Human T Cells

The piggyBac transposon system is naturally active, originally derived from the cabbage looper moth1,2. This non-viral system is plasmid based, most commonly utilizing two plasmids with one expressing the piggyBac transposase enzyme and a transposon plasmid harboring the gene(s) of interest between inverted repeat elements which are required for gene transfer activity. PiggyBac mediates gene transfer through a "cut and paste" mechanism whereby the transposase integrates the transposon segment into the genome of the target cell(s) of interest. PiggyBac has demonstrated efficient gene delivery activity in a wide variety of insect1,2, mammalian3-5, and human cells6 including primary human T cells7,8. Recently, a hyperactive piggyBac transposase was generated improving gene transfer efficiency9,10.
Human T lymphocytes are of clinical interest for adoptive immunotherapy of cancer11. Of note, the first clinical trial involving transposon modification of human T cells using the Sleeping beauty transposon system has been approved12. We have previously evaluated the utility of piggyBac as a non-viral methodology for genetic modification of human T cells. We found piggyBac to be efficient in genetic modification of human T cells with a reporter gene and a non-immunogenic inducible suicide gene7. Analysis of genomic integration sites revealed a lack of preference for integration into or near known proto-oncogenes13. We used piggyBac to gene-modify cytotoxic T lymphocytes to carry a chimeric antigen receptor directed against the tumor antigen HER2, and found that gene-modified T cells mediated targeted killing of HER2-positive tumor cells in vitro and in vivo in an orthotopic mouse model14. We have also used piggyBac to generate human T cells resistant to rapamycin, which should be useful in cancer therapies where rapamycin is utilized15.
Herein, we describe a method for using piggyBac to genetically modify primary human T cells. This includes isolation of peripheral blood mononuclear cells (PBMCs) from human blood followed by culture, gene modification, and activation of T cells. For the purpose of this report, T cells were modified with a reporter gene (eGFP) for analysis and quantification of gene expression by flow cytometry.
PiggyBac can be used to modify human T cells with a variety of genes of interest. Although we have used piggyBac to direct T cells to tumor antigens14, we have also used piggyBac to add an inducible safety switch in order to eliminate gene modified cells if needed7. The large cargo capacity of piggyBac has also enabled gene transfer of a large rapamycin resistant mTOR molecule (15 kb)15. Therefore, we present a non-viral methodology for stable gene-modification of primary human T cells for a wide variety of purposes.

piggyBac Transposon System Modification of Primary Human T Cells

We describe a method to genetically modify primary human T cells with a transgene using the non-viral piggyBac transposon system. T cells modified to using the piggyBac transposon system exhibit stable transgene expression.

The piggyBac transposon system is naturally active, originally derived from the cabbage looper moth1,2. This non-viral system is plasmid based, most ...

Isolation of Myeloid Dendritic Cells and Epithelial Cells from Human Thymus

In this protocol we provide a method to isolate dendritic cells (DC) and epithelial cells (TEC) from the human thymus. DC and TEC are the major antigen presenting cell (APC) types found in a normal thymus and it is well established that they play distinct roles during thymic selection. These cells are localized in distinct microenvironments in the thymus and each APC type makes up only a minor population of cells. To further understand the biology of these cell types, characterization of these cell populations is highly desirable but due to their low frequency, isolation of any of these cell types requires an efficient and reproducible procedure. This protocol details a method to obtain cells suitable for characterization of diverse cellular properties. Thymic tissue is mechanically disrupted and after different steps of enzymatic digestion, the resulting cell suspension is enriched using a Percoll density centrifugation step. For isolation of myeloid DC (CD11c+), cells from the low-density fraction (LDF) are immunoselected by magnetic cell sorting. Enrichment of TEC populations (mTEC, cTEC) is achieved by depletion of hematopoietic (CD45hi) cells from the low-density Percoll cell fraction allowing their subsequent isolation via fluorescence activated cell sorting (FACS) using specific cell markers. The isolated cells can be used for different downstream applications.

This protocol details a method to isolate antigen presenting cells from human thymus via different steps of enzymatic digestion of the tissue followed by density centrifugation of the single cell suspension and finally magnetic and/or FACS sorting of the cell populations of interest.

In this protocol we provide a method to isolate dendritic cells (DC) and epithelial cells (TEC) from the human thymus. DC and TEC are the major antigen ...

Assessing the Innate Sensing of HIV-1 Infected CD4+ T Cells by Plasmacytoid Dendritic Cells Using an Ex vivo Co-culture System.

HIV-1 innate sensing requires direct contact of infected CD4+ T cells with plasmacytoid dendritic cells (pDCs). In order to study this process, the protocols described here use freshly isolated human peripheral blood mononuclear cells (PBMCs) or plasmacytoid dendritic cells (pDCs) to sense infections in either T cell line (MT4) or heterologous primary CD4+ T cells. In order to ensure proper sensing, it is essential that PBMC are isolated immediately after blood collection and that optimal percentage of infected T cells are used. Furthermore, multi-parametric flow cytometric staining can be used to confirm that PBMC samples contain the different cell lineages at physiological ratios. A number of controls can also be included to evaluate viability and functionality of pDCs. These include, the presence of specific surface markers, assessing cellular responses to known agonist of Toll-Like Receptors (TLR) pathways, and confirming a lack of spontaneous type-I interferon (IFN) production. In this system, freshly isolated PBMCs or pDCs are co-cultured with HIV-1 infected cells in 96 well plates for 18-22 hr. Supernatants from these co-cultures are then used to determine the levels of bioactive type-I IFNs by monitoring the activation of the ISGF3 pathway in HEK-Blue IFN-&#945;/&#946; cells. Prior and during co-culture conditions, target cells can be subjected to flow cytometric analysis to determine a number of parameters, including the percentage of infected cells, levels of specific surface markers, and differential killing of infected cells. Although, these protocols were initially developed to follow type-I IFN production, they could potentially be used to study other imuno-modulatory molecules released from pDCs and to gain further insight into the molecular mechanisms governing HIV-1 innate sensing.

Assessing the Innate Sensing of HIV-1 Infected CD4+ T Cells by Plasmacytoid Dendritic Cells Using an Ex vivo Co-culture System.

Unlike cell-free HIV-1 particles, infected CD4+ T cells are effectively sensed by plasmacytoid dendritic cells (pDCs). This manuscript describes a method where peripheral blood mononuclear cells (PBMCs) or isolated pDCs are co-cultured with HIV-1 infected T cells to evaluate innate sensing by pDCs as assessed by release of type-I IFN.

HIV-1 innate sensing requires direct contact of infected CD4+ T cells with plasmacytoid dendritic cells (pDCs). In order to study this process, the ...

Generation of Human Alloantigen-specific T Cells from Peripheral Blood

The study of human T lymphocyte biology often involves examination of responses to activating ligands. T cells recognize and respond to processed peptide antigens presented by MHC (human ortholog HLA) molecules through the T cell receptor (TCR) in a highly sensitive and specific manner. While the primary function of T cells is to mediate protective immune responses to foreign antigens presented by self-MHC, T cells respond robustly to antigenic differences in allogeneic tissues. T cell responses to alloantigens can be described as either direct or indirect alloreactivity. In alloreactivity, the T cell responds through highly specific recognition of both the presented peptide and the MHC molecule. The robust oligoclonal response of T cells to allogeneic stimulation reflects the large number of potentially stimulatory alloantigens present in allogeneic tissues. While the breadth of alloreactive T cell responses is an important factor in initiating and mediating the pathology associated with biologically-relevant alloreactive responses such as graft versus host disease and allograft rejection, it can preclude analysis of T cell responses to allogeneic ligands. To this end, this protocol describes a method for generating alloreactive T cells from naive human peripheral blood leukocytes (PBL) that respond to known peptide-MHC (pMHC) alloantigens. The protocol applies pMHC multimer labeling, magnetic bead enrichment and flow cytometry to single cell in vitro culture methods for the generation of alloantigen-specific T cell clones. This enables studies of the biochemistry and function of T cells responding to allogeneic stimulation.

This article describes a method for the generation and propagation of human T cell clones that specifically respond to a defined alloantigen. This protocol can be adapted for cloning human T cells specific for a variety of peptide-MHC ligands.

The study of human T lymphocyte biology often involves examination of responses to activating ligands. T cells recognize and respond to processed peptide ...

Generation of CAR T Cells for Adoptive Therapy in the Context of Glioblastoma Standard of Care

Adoptive T cell immunotherapy offers a promising strategy for specifically targeting and eliminating malignant gliomas. T cells can be engineered ex vivo to express chimeric antigen receptors specific for glioma antigens (CAR T cells). The expansion and function of adoptively transferred CAR T cells can be potentiated by the lymphodepletive and tumoricidal effects of standard of care chemotherapy and radiotherapy. We describe a method for generating CAR T cells targeting EGFRvIII, a glioma-specific antigen, and evaluating their efficacy when combined with a murine model of glioblastoma standard of care. T cells are engineered by transduction with a retroviral vector containing the anti-EGFRvIII CAR gene. Tumor-bearing animals are subjected to host conditioning by a course of temozolomide and whole brain irradiation at dose regimens designed to model clinical standard of care. CAR T cells are then delivered intravenously to primed hosts. This method can be used to evaluate the antitumor efficacy of CAR T cells in the context of standard of care.

The lymphodepletive and immunomodulatory effects of chemotherapy and radiation standard of care can be leveraged to enhance the antitumor efficacy of T cell immunotherapy. We outline a method for generating EGFRvIII-specific chimeric antigen receptor (CAR) T cells and administering them in the context of glioblastoma standard of care.

Adoptive T cell immunotherapy offers a promising strategy for specifically targeting and eliminating malignant gliomas. T cells can be engineered ex vivo ...

Assessment of the Immunomodulatory Properties of Human Mesenchymal Stem Cells (MSCs)

The immunomodulatory properties of multilineage human mesenchymal stem cells (MSCs) appear to be highly relevant for clinical use towards a wide-range of immune-related diseases. Mechanisms involved are increasingly being elucidated and in this article, we describe the basic experiment to assess MSC immunomodulation by assaying for suppression of effector leukocyte proliferation. Representing activation, leukocyte proliferation can be assessed by a number of techniques, and we describe in this protocol the use of the fluorescent cellular dye carboxyfluorescein succinimidyl ester (CFSE) to label leukocytes with subsequent flow cytometric analyses. This technique can not only assess proliferation without radioactivity, but also the number of cell divisions that have occurred as well as allowing for identification of the specific population of proliferating cells and intracellular cytokine/factor expression. Moreover, the assay can be tailored to evaluate specific populations of effector leukocytes by magnetic bead surface marker selection of single peripheral blood mononuclear cell populations prior to co-culture with MSCs. The flexibility of this co-culture assay is useful for investigating cellular interactions between MSCs and leukocytes.

Assessment of the Immunomodulatory Properties of Human Mesenchymal Stem Cells MSCs

The immunomodulatory properties of human mesenchymal stem cells (MSC) appear increasingly relevant for clinical application. Using a co-culture system of MSCs and peripheral blood leukocytes pre-stained with the fluorescent dye carboxyfluorescein succinimidyl ester (CFSE), we describe the in vitro assessment of MSC immunomodulation on effector leukocyte proliferation and specific subpopulations.

The immunomodulatory properties of multilineage human mesenchymal stem cells (MSCs) appear to be highly relevant for clinical use towards a wide-range of ...

A Novel Feeder-free System for Mass Production of Murine Natural Killer Cells In Vitro

Natural killer (NK) cells belong to the innate immune system and are a first-line anti-cancer immune defense; however, they are suppressed in the tumor microenvironment and the underlying mechanism is still largely unknown. The lack of a consistent and reliable source of NK cells limits the research progress of NK cell immunity. Here, we report an in vitro system that can provide high quality and quantity of bone marrow-derived murine NK cells under a feeder-free condition. More importantly, we also demonstrate that siRNA-mediated gene silencing successfully inhibits the E4bp4-dependent NK cell maturation by using this system. Thus, this novel in vitro NK cell differentiating system is a biomaterial solution for immunity research.

A Novel Feeder-free System for Mass Production of Murine Natural Killer Cells In Vitro

Here, we present a protocol to mass-produce gene-silencing murine NK cells by using a feeder-free differentiation system for mechanistic study in vitro and in vivo.

Natural killer (NK) cells belong to the innate immune system and are a first-line anti-cancer immune defense; however, they are suppressed in the tumor ...

Development and Validation of an Ultrasensitive Single Molecule Array Digital Enzyme-linked Immunosorbent Assay for Human Interferon-&#945;

The main aim of this protocol is to describe the development and validation of an interferon (IFN)-&#945; single molecule array digital Enzyme-Linked ImmunoSorbent Assay (ELISA) assay. This system enables the quantification of human IFN-&#945; protein with unprecedented sensitivity, and with no cross-reactivity for other species of IFN.
The first key step of the protocol is the choice of the antibody pair, followed by the conjugation of the capture antibody to paramagnetic beads, and biotinylation of the detection antibody. Following this step, different parameters such as assay configuration, detector antibody concentration, and buffer composition can be modified until optimum sensitivity is achieved. Finally, specificity and reproducibility of the method are assessed to ensure confidence in the results. Here, we developed an IFN-&#945; single molecule array assay with a limit of detection of 0.69 fg/mL using high-affinity autoantibodies isolated from patients with biallelic mutations in the autoimmune regulator (AIRE) protein causing autoimmune polyendocrinopathy syndrome type 1/autoimmune polyendocrinopathy-candidiasis-ectodermal dystrophy (APS1/APECED). Importantly, these antibodies enabled detection of all 13 IFN-&#945; subtypes.
This new methodology allows the detection and quantification of IFN-&#945; protein in human biological samples at attomolar concentrations for the first time. Such a tool will be highly useful in monitoring the levels of this cytokine in human health and disease states, most particularly infection, autoimmunity, and autoinflammation.

Development and Validation of an Ultrasensitive Single Molecule Array Digital Enzyme-linked Immunosorbent Assay for Human Interferon-&amp;#945;

Here we present a protocol to describe the development and validation of a single molecule array digital ELISA assay, which enables the ultra-sensitive detection of all IFN-&#945; subtypes in human samples.

The main aim of this protocol is to describe the development and validation of an interferon (IFN)-&#945; single molecule array digital Enzyme-Linked ...